1. Preparation of target-specific primer working stocks


For multiplexed Illumina sequencing-by-synthesis, each sample must be uniquely tagged with molecular barcodes to extract the reads per sample out of the pool of sequence reads. These tags are added using a 4-primer PCR system developed by Fluidigm. We order fusion primers containing our MHC class I and class II DRB sequence-specific PCR primers prefaced with Fluidigm Consensus Sequence (CS) linkers. Barcodes are then added via commercially available plates of outer primers ordered from Fluidigm that contain the complements to the CS linkers on 384 different barcode sequences.

Our MHC-I exon 2 primer (SBT195) is staggered (the blue nucleotides below) to ensure amplicon diversity during the sequencing run.
  Primer Name Linker Sequence (5'-3') MHC-Specific Sequence (5'-3')
Pair 1 CS1-SBT195R ACACTGACGACATGGTTCTACA GCCTCGCTCTGGTTGTAGTAG
  CS2tca-SBT195F TACGGTAGCAGAGACTTGGTCT TCAGGGCTACGTGGACGACAC
Pair 2 CS1t-SBT195R ACACTGACGACATGGTTCTACA TGCCTCGCTCTGGTTGTAGTAG
  CS2ca-SBT195F TACGGTAGCAGAGACTTGGTCT CAGGGCTACGTGGACGACAC
Pair 3 CS1ct-SBT195R ACACTGACGACATGGTTCTACA CTGCCTCGCTCTGGTTGTAGTAG
  CS2a-SBT195F TACGGTAGCAGAGACTTGGTCT AGGGCTACGTGGACGACAC
Pair 4 CS1act-SBT195R ACACTGACGACATGGTTCTACA ACTGCCTCGCTCTGGTTGTAGTAG
  CS2-SBT195F TACGGTAGCAGAGACTTGGTCT GGGCTACGTGGACGACAC

In our experience, only these antisense SBT195 primer pairs consistently amplify the desired product. The CS1-F and CS2-R primers did not work as consistently.

Our DRB283g primer amplifies DRB exon 2 reads from gDNA. We have not incorporated staggering since we generally sequence DRB along with other amplicons. Using both the sense and antisense versions will improve diversity somewhat.
  Primer Name Linker Sequence (5'-3') MHC-Specific Sequence (5'-3')
Pair 1 (antisense) CS1-DRB283gR ACACTGACGACATGGTTCTACA CCTCGCCGCTGCACTGT
  CS2-DRB283gF TACGGTAGCAGAGACTTGGTCT TCCCCACAGCACGTTTCTT
Pair 2 (sense) CS1-DRB283gF ACACTGACGACATGGTTCTACA TCCCCACAGCACGTTTCTT
  CS2-DRB283gR TACGGTAGCAGAGACTTGGTCT CCTCGCCGCTGCACTGT

Our DRB283 amplicon is similar to the DRB283g primers, but the forward primers lack intronic sequence; they instead span into exon 1 and will thus only amplify cDNA templates robustly. We again use the antisense versions with staggering (like the SBT195 primers).
  Primer Name Linker Sequence (5'-3') MHC-Specific Sequence (5'-3')
Pair 1 CS1-DRB283R ACACTGACGACATGGTTCTACA ACTCGCCGCTGCACTGT
  CS2ggt-DRB283F TACGGTAGCAGAGACTTGGTCT GGTACACCCGACCACGTTTCTT
Pair 2 CS1t-DRB283R ACACTGACGACATGGTTCTACA TACTCGCCGCTGCACTGT
  CS2ct-DRB283F TACGGTAGCAGAGACTTGGTCT CTACACCCGACCACGTTTCTT
Pair 3 CS1ct-DRB283R ACACTGACGACATGGTTCTACA CTACTCGCCGCTGCACTGT
  CS2t-DRB283F TACGGTAGCAGAGACTTGGTCT TACACCCGACCACGTTTCTT
Pair 4 CS1tgt-DRB283R ACACTGACGACATGGTTCTACA TGTACTCGCCGCTGCACTGT
  CS2-DRB283F TACGGTAGCAGAGACTTGGTCT ACACCCGACCACGTTTCTT

2. PCR setup and thermocycling conditions


(a) Normalize gDNA/cDNA to ~10 ng/µl. Templates within 4-fold of this concentration will work.
(b) Make PCR master mix. For each sample (+3x overage):
   12.5 µl 2X Phusion High Fidelity PCR Master Mix (New England Biolabs)
   7.5 µl PCR grade H2O
(c) In each well, combine 20 µl PCR master mix with 2.5 µl of 0.125 µM target-specific primer working stock, 1.25 µl of 2 µM Fluidigm barcoded Illumina adaptor primers and 1 µl of gDNA/cDNA template.
(d) Amplify on thermocycler using the program below, using 30 cycles for gDNA and 25 cycles for cDNA.
98°C 3 min - denaturation + polymerase activation
30 / 25 cycles of:
 98°C 5 s - denaturation
 60°C 10 s - annealing
 72°C 20 s - extension
72°C 5 min - final extension

3. Confirmation of PCR products by FlashGel (Lonza)


(a) Open FlashGel package, and carefully pour a few drops of water into wells, tilt cartridge until water fills each well, pour off any excess.
(b) Into an empty 96-well plate or set of strip tubes, pipet:
   1 µl 5X loading dye
   4 µl PCR product
(c) Pipet up and down 5 times to mix.
(d) Add 5 µl of sample to each well, reserving the smaller leftmost well.
(e) Add 5 µl of QuantLadder to leftmost well.
(f) Fully attach the cassette to the electrodes and run at 270 mV until ladder has spread out.
(g) Image gel on imager:
   For SBT195, you are looking for a band just above the 250 bp marker (2nd band)
   For DRB283 primers, it is just above the 400 bp marker (3rd band)
(h) If a band is not visible for a sample, you may decide to give it 5 extra thermal cycles.

4. Ampure XP (Agencourt) bead purification of PCR products


We use two rounds of bead cleanup to remove short product contaminants from our amplicons:

(a) Calculate the volume of remaining sample in each tube/plate. Into an empty 96-well, 200 µl skirted plate, pipet 25 µl (~1.3X volume of sample) of Ampure XP beads. Add 20 µl sample and pipet up and down ten times to mix, trying to minimize bubbles.
(b) Incubate for 5 min at room temperature.
(c) Place plate on a six-bar magnet stand for 2 min.
(d) Being careful to avoid disrupting the bead pellets, remove all supernatant and discard.
(e) Rinse bead pellets with 180 µl 70% ethanol.
(f) Incubate for 30 s, discard supernatant.
(g) Repeat (f) for a second wash with 180 µl of 70% ethanol.
(h) After completely removing supernatant, remove plate from magnet and allow 2-4 min to dry (it's okay if it doesn't dry completely).
(i) Add 20 µl 1X TE buffer to each well to elute purified product and pipet mix 10 times (use 25 µl 1X TE buffer for the second elution).
(j) Incubate for 3 min.
(k) Place plate on magnet for 2 min (on second clean-up, proceed to step (n) after beads are pelleted).
(l) Pipet (20 µl*1.3=) 25 µl of Ampure XP beads into wells of a new 96-well 200 µl skirted plate. Transfer the eluted, purified product from the first plate into these new wells and pipet mix 10 times.
(m) Repeat steps (b)-(k) for second clean-up.
(n) Transfer eluted products to a new 96-well plate for storage/pooling.


5. Quantification of amplicon concentration with PicoGreen (Invitrogen) on a plate reader


(a) Prepare 8 DNA standards using serial 2:1 dilutions from 10 ng/µl DNA Standard in water in a strip of PCR tubes.
Concentration of standards should be as follows (in ng/ml):
   10
   5
   2.5
   1.25
   0.625
   0.312
   0.156
   0.078
(b) Each plate can quantify a maximum of 88 samples. Use multiple plates, if required.
(c) Make a master mix containing:
   199 µl 1X TE * (# of samples + 8 standards/plate)
   1 µl PicoGreen fluorescent dye * (# of samples + 8 standards/plate)
(d) Vortex and aliquot 195 µl of master mix into (# of samples/plate + 8 standards) wells of a black Fluotrac 200 plate (Greiner).
(d) Add 5 µl of sample into each sample well, and 5 µl of the standards into each standard well.
(e) Attach a clear, optical film seal to the plate and tighten with rubber spatula.
(e) Vortex the plate for 5 s. Spin the plate at 930 rcf for 1 min.
(f) Put the plate in the plate reader, use the Multimode Analysis software (Beckman Coulter) to enter the layout of samples/standards in the plate, enter the concentrations of each standard, and run the machine.
(g) Check the standard curve as a linear fit, and if satisfied save the resulting Excel output containing sample concentrations to a flashdrive.

6. Pooling of samples and library preparation for Illumina MiSeq sequencing


(a) Using the platereader output, select an amount of sample (in ng) that can be easily pipetted (between 0.5 µl and 20 µl) for as many samples as possible.
(b) Pipet the selected quantity of each sample into a single 1.5 ml tube.
(c) We perform two additional Ampure XP bead cleanups on this pool to remove any remaining dimers:

Calculate the volume of the pool, and confirm by drawing the expected volume into a pipet tip. Into the tube containing the pool, pipet 1.3X (volume of pool) of Ampure XP beads. Vortex briefly to mix.

Incubate for 5 min at room temperature.
Place tube on a tube magnet stand for 2 min.
Being careful to avoid disrupting the bead pellets, remove all supernatant and discard.
Rinse bead pellets with 950 µl 70% ethanol.
Incubate for 30 s, discard supernatant.
Repeat for a second wash with 950 µl of 70% ethanol.
After completely removing supernatant, remove tube from magnet and allow 2-4 min to dry (it's okay if it doesn't dry completely).
Add 100 µl 1X TE buffer to elute purified product and vortex briefly to mix. OPTIONAL: during the second elution, you can concentrate the pool by eluting in <100 µl 1X TE. This is useful when one pool is expected to be much less concentrated than other pools in the same MiSeq run.
Incubate for 3 min.
Place tube on magnet for 2 min (on second clean-up, skip to after beads are pelleted).
Pipet (100 µl*1.3=) 130 µl of Ampure XP beads into wells of a new 1.5 ml tube. Transfer the eluted, purified product from the first tube into this tube and vortex briefly to mix.
Return to ★ for second clean-up.
Transfer eluted products to a new 1.5 ml tube for storage/sequencing.
(d) We now quantify the pool using the Qubit dsDNA High Sensitivity Assay Kit (Life Technologies)

Make a master mix containing:
199 µl Qubit HS buffer * (# of samples + 2 standards)
1 µl Qubit HS dye * (# of samples + 2 standards)
Vortex and aliquot 190 µl of master mix into two Qubit tubes and 198 µl of master mix into each sample Qubit tube.
Add 2 µl of sample into each sample tube, and 10 µl of the standards (10 ng/µl and 0 ng/µl) into the standard tubes.
Vortex the tubes for 5 s. Tap/flick tubes to allow any bubbles to escape.
Measure the concentration of each tube in a Qubit fluorimeter. Convert to sample concentrations multiplying by the conversion factor 0.1 (as per the this calculation):
conc (ng/ml) * 200 µl/2 µl * 1 ml/1000 µl = conc * 0.1 ng/µl

7. Sequencing on the Illumina MiSeq


(a) Thaw a MiSeq reagent cartridge at RT.
(b) Generate a 2 nM stock of each sample using the Illumina copy number spreadsheet, along with the Qubit concentration and a DNA fragment size of 386bp.
(c) Pipet 2 µl of 2 nM stock from each sample into a 1.5 ml tube to create an undiluted amplicon pool.
(d) Pipet 10 µl of the DNA pool into a fresh 1.5 ml tube. Add 10 µl of 0.2 N NaOH solution to the DNA.
(e) Vortex briefly, then centrifuge for 1 min at 280 x g.
(f) Incubate for 5 min at RT to denature the DNA.
(g) Add 980 µl pre-chilled HT1 to the denatured DNA to generate a 20 pM denatured library.
(h) Transfer 600 µl of the denatured library to a new 1.5 ml tube, then add 400 µl of pre-chilled HT1 to dilute the denatured library to 12 pM.
(i) Invert several times to mix, then centrifuge briefly to get any droplets off of the cap. Place on ice.
(j) Pipet 2 µl of 10 nM PhiX library into a fresh 1.5 ml tube. Add 8 µl of 10 mM Tris-Cl (pH 8.5 with 0.1% Tween 20) to create a 2 nM dilution of PhiX library. PhiX is used as an internal control for sequencing.
(k) Combine 10 µl of 2 nM PhiX library with 10 µl of 0.2 N NaOH solution in a 1.5 ml tube to create a 1 nM PhiX library.
(l) Vortex briefly, then centrifuge for 1 min at 280 x g.
(m) Incubate for 5 min at RT to denature the PhiX library.
(n) Add 980 µl pre-chilled HT1 to the denatured PhiX library to generate a 20 pM PhiX library. Note: The denatured 20 pM PhiX library can be stored up to 3 weeks at -20°C.
(o) Transfer 12 µl of 20 pM PhiX library to a new 1.5 ml tube, then add 8 µl of pre-chilled HT1 to dilute the PhiX library to 12 pM.
(p) Combine 990 µl of denatured library (12 pM) and 10 µl of PhiX library (12 pM) to create a final pool for sequencing. Place on ice.
(q) Use an empty 1 ml pipet tip to pierce the foil seal of the MiSeq reagent cartridge over the reservoir labeled “Load Samples”.
(r) Pipet 600 µl of the final pool from step 16 into the “Load Samples” reservoir. Avoid touching the foil seal while dispensing the sample.
(s) Follow the onscreen prompts on the Illumina MiSeq instrument to set up and perform the sequencing run.